Seeding 
After cells are thawed:

NOTE: Do not dispense the entire contents of the cryovial into one T-25 flask!!

1. Remove the cap, being careful not to touch the interior threads with your fingers.

2. Using a micro-pipette with a 1000 ml tip set to 800 ml, put the tip into the cryovial and resuspend the cells, with a gentle, slow and steady up and down pipetting motion no more than five times. DO NOT resuspend quickly, and keep the tip near the bottom to avoid making bubbles.

3. Dispense an equal amount of cells into the flasks. If five T-25 flasks were prepared, set micropipetter to 200 ml and dispense.

4. Replace the cap or cover, and gently rock the vessels to evenly distribute the cells. Loosen caps if necessary to permit gas exchange (see "Set Up," step number 4, pg. 11).

5. Return the culture vessels to 37·C, 5% CO2 incubator. Lay them flat on the shelf, providing the largest surface for cells to attach. The cells will anchor to the bottom surface of the flask.

Maintenance After Seeding
Normal Human Fibroblasts are not tolerant of rapid temperature fluctuations or nutrient-deficient medium. Feeding them with fresh growth medium that has been warmed will avert potential problems. (Remember to warm only the amount needed.) Check and feed the cells on the schedule below, even on weekends and holidays.

1. Change the growth medium the day after seeding (to remove residual DMSO and unattached cells), then every other day thereafter while examining them daily.

NOTE: A change of medium requires removal of the medium by aspirating with a sterile pipette on the opposite side of the flask from where the cells are attached. Then warm, fresh medium is added down that same side.

2. Successfully recovered cultures will exhibit the following:

a. Cells with clear non-granular cytoplasm.

b. Numerous mitotic figures after day 2.

3. Feed the cells a larger volume of medium as they become more confluent. Use this table as a guideline:

4. Continue feeding the cells until 70 - 90% confluence. If the cells are allowed to become over-confluent they will suffer contact inhibition and will pop off the flask and/or be difficult to trypsinize.

Overview of Subculture Preparation

Subculture Preparation 
NOTE: The following instructions are for a 25 cm2 flask. Adjust all volumes accordingly Preparation for other size flasks.

Preparation for subculturing the first flask:

1. Subculture the cells when they are 70-90% confluent and contain many mitotic figures throughout the flask.

2. For each 25 cm2 of cells to be subcultured, allow 3 ml of Clonetics® Trypsin/EDTA (T/E) to thaw and come to room temperature. For NHDF grown in FGM® use cold T/E (4·C) for subculturing.

3. For each 25 cm2 of cells to be subcultured, allow 5 ml of Clonetics® HEPES Buffered Saline Solution (HEPES-BSS) to come to room temperature.

4. For each 25 cm2 of cells to be subcultured, allow 3 ml of Trypsin Neutralizing Solution (TNS) to come to room temperature.

5. Remove growth medium from 4·C storage and allow to start warming to room temperature.

6. Have flasks available for seeding cells.

Subculturing 
Subculture one flask at a time. All flasks following the first flask will be subcultured following an optimization of this protocol (explained later in this procedure), based on calculated cell count, cell viability, and seeding density.

In a sterile field:

1. Aspirate the medium from one culture vessel.

2. For NHLF and NHDF grown in FGM®-2, rinse the cells with 2 - 3 ml of room temperature HEPES-BSS. DON'T forget this step. The medium contains complex proteins that neutralize the trypsin, making it ineffective. For NHDF grown in FGM®, skip steps 2 and 3.

3. Aspirate the HEPES-BSS from the flask.

4. Cover the cells with 3 ml of Clonetics® T/E solution. Use cold (4·C) T/E solution for NHDF grown in FGM®.

5. Rock the flask to make sure all cells come into contact with the trypsin.

6. Tighten the cap and begin monitoring the flask under the microscope.

7. Continue to examine the cell layer microscopically. 
   a. Allow the trypsinization to continue until 3 90% of the cells are rounded up.
   
   NOTE: Rounded up cells are spherical, have smooth edges and are refractile or shiny. If the cells still have protruding nubs which are still attached to the flask, they need more time to trypsinize. This entire process takes about 1 to 2 minutes, under optimal conditions.
   
   b. At this point, rap the flask against the palm of your hand to release the majority of cells from the culture surface. If only a few cells detach, you may not have let them trypsinize long enough. Wait 30 seconds and rap again. If cells still do not detach, wait and rap every 30 seconds thereafter. 
   
   NOTE: Don't try to get all cells to detach by rapping them severely. This action may damage the cells.
   

8. After cells are released, neutralize the trypsin in the flask with 3 ml of room temperature TNS.
   
   If the majority of cells do not detach within four minutes, the trypsin is either not warm enough or not active enough to release the cells. Harvest the culture vessel as described above, and either re-trypsinize with fresh, warm Clonetics® Trypsin/EDTA Solution (or) rinse with Clonetics® Trypsin Neutralizing Solution and then add fresh, warm growth medium to the culture vessel and return to an incubator until fresh trypsinization reagents are available.
   

9. Quickly transfer the detached cells to a sterile 15 ml centrifuge tube.

10. Rinse the flask with a final 2 ml of HEPES-BSS to collect residual cells, and add this rinse to the centrifuge tube.

11. Examine the harvested flask under the microscope to make sure the harvest was successful by looking at the number of cells left behind. This should be less than 5%.

12. Centrifuge the harvested cells at 220 x g for 5 minutes to pellet the cells. 
    a. Aspirate most of the supernatant, except for 100-200 ml.
    b. Flick the cryovial with your finger to loosen the pellet.
    

13. Dilute the cells in 4-5 ml of growth medium and note the total volume of the diluted cell suspension.

14. During these procedures keep the cells in ice until they are plated.
    
    To obtain the best results from your cells, you will assess cell yield and viability with Trypan Blue. Trypan Blue is a dye used to highlight dead cells. Dead cells take up the dye and appear blue, instead of refractile and colorless. Evaluate on bright-field microscope. Follow these steps:
    

15. Count the cells with a hemacytometer or cell counter and calculate the total number of cells. (See Appendix B.) Make a note of your cell yield for later use.
    
    The cell suspension should contain between 250,000 to 1,000,000 cells/ml for greatest accuracy.
    

16. If necessary, dilute the suspension with HEPES Buffered Saline Solution (HEPES-BSS) to achieve the desired "cells/ml" and re-count the cells.

17. Assess cell viability using Trypan Blue (see Appendix C).

18. Use the following equation to determine the total number of viable cells:
    
    Total # of Viable Cells = Total cell count x percent viability / 100
    
    Example: 1,000,000 cells x 60 / 100 = 600,000 viable cells 
    

19. Determine the total number of flasks to inoculate by using the following equation. The number of flasks needed depends upon cell yield and seeding density. Larger flasks may be used to save plasticware and time spent on subsequent subcultures. Smaller flasks reduce the risk of losing a substantial part of your culture if contamination occurs.
    
    NOTE: Recommended seeding density is 3500 cells/cm2 for NHDF and 2500 cells/cm2 for NHLF.
    
    Total # of flasks to inoculate = Total # of viable cells / Growth Area of Flask x Recommended Seeding Density
    
    Example:600,000 viable cells / 75 cm2 x 3500 cells/cm2 = 2 T-75 flasks (rounded down to nearest whole number)
    

20. Use the following equation to calculate the volume of cell suspension to seed into your flasks.
    
    Seeding volume = Total volume of diluted cell suspension / # of flasks as determined in step 18
    
    Example: 4.3 ml of diluted cell suspension / 2 T-75 flasks = 2.15 ml per T-75 flask
    

21. Prepare flasks by labeling each flask with the passage number, strain number, cell type, and date.

22. Carefully open the medium bottle and transfer growth medium to new culture vessels by adding 1 ml growth medium for every 5 cm2 surface area of the flask (1ml/5cm2). 
    
    Example:15 ml growth medium for a 75 cm2 flask.
    

23. After mixing the diluted cells with a 5 ml pipet to ensure a uniform suspension, dispense the volume of suspension calculated above into the prepared subculture flasks.

24. After dispensing the cells, gently rock flask to promote even distribution.

25. If not using vented caps, loosen caps of flasks. Place the new culture vessels into a 37·C humidified incubator with 5% CO2..

Assessing Yield and Viability
Several factors contribute to low cell count and low cell viability. An example of yield viability assessment is provided in the chart below. To determine the reason for low yield/viability, follow these steps:

1. Study the sample chart below. It is a sample of high yield, high viability.

a. Note the "solid dot" on the far, left side of the square. It indicates high yield, or a cell count of more than 250,000 for NHDF and more than 500,000 for NHLF.

b. Note the "solid dot" on the X axis or bottom line of the square. It indicates high viability, or more than 50% viability.

c. Extend a line from each dot as shown in the chart. The point where the lines intersect (the bold "X") is located in the High Yield/High Viability quadrant. Thus, the sample is optimal.

2. Now, using the blank diagram below plot your cell yield and cell viability. Follow these steps:

a. Mark a (·) on the Y axis to indicate the total cell count of your culture.

b Mark a (·) on the X axis to indicate the calculated percent viability of your culture.